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Abstract

Background

The oxidation of carbohydrates from lignocellulose can facilitate the synthesis of
new biopolymers and biochemicals, and also reduce sugar metabolism by lignocellulolytic
microorganisms, reserving aldonates for fermentation to biofuels. Although oxidoreductases
that oxidize cellulosic hydrolysates have been well characterized, none have been
reported to oxidize substituted or branched xylo-oligosaccharides. Moreover, this
is the first report that identifies amino acid substitutions leading to GOOX variants
with reduced substrate inhibition.

Results

The recombinant wild type gluco-oligosaccharide oxidase (GOOX) from the fungus Sarocladium strictum, along with variants that were generated by site-directed mutagenesis, retained the
FAD cofactor, and showed high activity on cello-oligosaccharide and xylo-oligosaccharides,
including substituted and branched xylo-oligosaccharides. Mass spectrometric analyses
confirmed that GOOX introduces one oxygen atom to oxidized products, and 1H NMR and tandem mass spectrometry analysis confirmed that oxidation was restricted
to the anomeric carbon. The A38V mutation, which is close to a predicted divalent
ion-binding site in the FAD-binding domain of GOOX but 30 Å away from the active site,
significantly increased the kcat and catalytic efficiency of the enzyme on all oligosaccharides. Eight amino acid
substitutions were separately introduced to the substrate-binding domain of GOOX-VN
(at positions Y72, E247, W351, Q353 and Q384). In all cases, the Km of the enzyme variant was higher than that of GOOX, supporting the role of corresponding
residues in substrate binding. Most notably, W351A increased Km values by up to two orders of magnitude while also increasing kcat up to 3-fold on cello- and xylo-oligosaccharides and showing no substrate inhibition.

Conclusions

This study provides further evidence that S. strictum GOOX has broader substrate specificity than the enzyme name implies, and that substrate
inhibition can be reduced by removing aromatic side chains in the -2 binding subsite.
Of the enzyme variants, W351A might be particularly advantageous when oxidizing oligosaccharides
present at high substrate concentrations often experienced in industrial processes.

Early reports of GOOX-T1 from the fungus Sarocladium strictum T1 (previously known as Acremonium strictum T1 [8]) confirmed oxidation of the hydroxyl group attached to the anomeric carbon of maltose
[6]; other analyses revealed even higher activities on cello-oligosaccharides, particularly
cellotriose [9,10]. Like other flavin carbohydrate oxidases that target the hydroxyl group of the anomeric
carbon, GOOX-T1 is thought to mediate oxidoreductase activity through two half-reactions:
1) oxidation of the reducing sugar to the corresponding lactone, and 2) reduction
of molecular oxygen to hydrogen peroxide [11]. Subsequent hydrolysis of the lactone product to the corresponding carboxylic acid
may then occur. While the biological function of GOOX is uncertain, hydrogen peroxide
generated through carbohydrate oxidation could be used by lignin peroxidases and manganese
peroxidase in lignin degradation. From an applied perspective, gluco-oligosaccharide
oxidases could provide an alternative to CDHs used in amperometric enzyme biosensors
for real-time measurement of cellulase activity on insoluble cellulose [12]. More recent applications of CDH also demonstrate the benefit of carbohydrate oxidation
to reduce sugar consumption by lignocellulolytic fungi, thereby maximizing ethanol
yields from fermenting microorganisms [13].

The crystal structure of GOOX-T1 reveals a monomeric glycoprotein with a flavin adenine
dinucleotide (FAD)-binding domain coordinated by a bi-covalent linkage to H70 (8α-N1-histidyl)
and C130 (6-S-cysteinyl); GOOX-T1 is also characterized by having a comparatively
open substrate-binding site [14]. Site-directed mutagenesis confirmed the requirement of bi-covalent coordination
of FAD for enzyme activity; this unique coordination is also correlated to the relatively
high redox potential of GOOX-T1 [14,15]. In our recent study, GOOX-VN from S. strictum strain CBS 346.70 was recombinantly expressed and biochemically characterized using
a range of sugars and oligosaccharides, including cello-oligosaccharides and xylo-oligosaccharides
with up to 3 sugar units [7]. Fifteen amino acid differences distinguish GOOX-VN and GOOX-T1: 13 are intrinsic
differences in the wild-type gene sequences while 2 (A38V and S388N) arose from random
mutations during the construction of the GOOX-VN expression system [7] (Additional file 1: Figure S1). GOOX-VN was found to oxidize xylose as well as xylobiose and xylotriose
[7]. Given the high sequence identity between GOOX-VN and GOOX-T1 (97%), and since none
of the amino acid substitutions between GOOX-VN and GOOX-T1 are predicted to directly
participate in substrate binding, it is likely that GOOX-T1 also oxidizes xylo-oligosaccharides
even though xylo-oligosaccharide oxidation by GOOX-T1 has not been reported [7,10]. Notably, resulting enzymatically oxidized oligosaccharides could be used as carbohydrate
standards that replaces the comparatively arduous chemical synthesis approach [16], facilitating the characterization of carbohydrate-oxidizing enzymes whose activity
can not be easily measured by colorimetric assays.

Additional file 1: Figure S1. Protein sequence alignment of GOOX-T1, GOOX and GOOX-VN. The protein sequence of GOOX-T1
from S. strictum T1 was aligned with those of GOOX and GOOX-VN from S. strictum CBS 346.70. The positions of amino acid differences are numbered while the amino
acid substitutions created in GOOX-VN for the current study are indicated by rectangles.
Amino acid substitutions introduced to re-construct GOOX from GOOX-VN are indicated
by an asterisk.

To investigate the role of selected amino acids on substrate preference, three amino
acids in the GOOX-VN substrate binding site were previously substituted to corresponding
residues in chito-oligosaccharide oxidase (ChitO) from Fusarium graminearum[15] or carbohydrate oxidase from Microdochium nivale[17], which show 45% and 42% sequence identity to GOOX-VN, respectively [7]. Of these, Y300A nearly doubled kcat values for oligosaccharides while also increasing corresponding Km values [7]. The current study describes a more comprehensive assessment of substrate preference
and catalysis by GOOX-VN by 1) constructing eight additional amino acid substitutions
within the substrate binding site of this enzyme, 2) generating V38A and N388S substitutions
that convert GOOX-VN to the wild-type GOOX sequence, and 3) using several oligosaccharides,
including branched xylo-oligosaccharides (Figure 1) to characterize the catalytic efficiency, substrate selectivity and substrate inhibition
of GOOX-VN enzyme variants. These analyses confirmed comparable kinetic efficiencies
on cello-oligosaccharides and xylo-oligosaccharides, suggesting that gluco-oligosaccharide
oxidases characterized to date have broader substrate specificity than the enzyme
name implies. This study also identified enzyme variants with high catalytic activity
but lower substrate inhibition, which could improve oligosaccharide oxidation at high
substrate concentrations often experienced in industrial bioprocesses.

Results and discussion

Protein expression and biophysical characterization

Recombinantly expressed GOOX-VN and enzyme variants were purified to more than 95%
homogeneity by affinity chromatography (Figure 2A, Additional file 2: Figure S2). Amino acid substitutions did not affect protein yields, and in general,
between 5 and 10 mg/L of purified protein were recovered. The observed mass of all
enzymes was approximately 70 kDa (Additional file 2: Figure S2), suggesting that glycosylation could account for approximately 20% of
the protein, which is similar to the mass percentage of carbohydrates in glucose oxidase
[18]. Notably, the deglycosylation of GOOX-VN by PNGaseF, generated a band at about 56
kDa on SDS-PAGE gels [7], but this deglycosylation did not affect the activity or substrate specificity of
the enzyme (Table 1).

Figure 2.The presence of the covalent FAD cofactor in GOOX. (A): An SDS-PAGE of GOOX and mutant variants stained by Coomassie blue (upper) and under
254 nm transillumination (lower), which shows intrinsic fluorescence of the covalently-bound
FAD upon acidification; protein samples were overloaded to facilitate the detection
of FAD intrinsic fluorescence. (B): UV–VIS scanning of GOOX, showing two automatically-determined maxima of 375 and
440 nm, which are similar to the absorbance (Abs) peaks of GOOX-T1 at 380 and 444
nm, respectively [10]; the 440-nm peak disappeared when the enzyme was reduced by 50 mM sodium hydrosulfite
(dotted solid line) or by 200 mM cellobiose (dash line).

Table 1.Effect of glycosylation on GOOX activity on cello- and xylo-oligosaccharides

None of the amino acid substitutions appeared to affect FAD binding, as assessed by
fluorescence detection (Figure 2A) and UV–VIS scanning (Figure 2B). Enhancement of fluorescence following performic acid oxidation is a convenient
method for detecting the presence of 8α-S-cysteinyl riboflavins [19]. Since pre-treatment of SDS-PAGE gels with performic acid did not increase the fluorescence
measured from GOOX-VN or any of the enzyme variants, one of the covalent linkages
to the FAD cofactor is likely 6-S-cysteinyl as seen in GOOX-T1 structures [14,20]. Moreover, because the flavinylation process is thought to promote proper protein
folding [14], detection of the FAD cofactor suggests that enzyme variants have assumed the correct
protein conformation.

Confirming the regioselectivity of gluco-oligosaccharide oxidases

To date, very few studies have confirmed the position of hydroxyl groups oxidized
by family AA7 gluco-oligosaccharide oxidases. Lee at al. [6] used 13C and 1H NMR to confirm that GOOX-T1 targets the hydroxyl group of the anomeric carbon, however,
only maltose was used in their analysis. Since gluco-oligosaccharide oxidase activity
is higher on cello-oligosaccharides and xylo-oligosaccharides than maltose [6,7], 1H NMR was used here to evaluate the effect of sugar type and linkage on the regio-selectivity
of GOOX enzymes.

The disappearance of H1 doublet signals from the reducing end of α- and β-glucose
units of cellobiose is consistent with oxidation at the anomeric C1 position (Figure 3A) [21]. Similarly, the peak height for the H1 signals from the reducing end of α- and β-xylose
units of xylobiose was decreased in oxidized xylobiose samples (Figure 3B). Ring opening at the anomeric position was also revealed by the detection of H2
and H3 signals at 4.05 ppm and 3.95 ppm in case of oxidized cellobiose, and at 4.01
ppm and 3.81 ppm, respectively in case of oxidized xylobiose [21,22]. The signals for the corresponding lactone were not observed probably due to the
relatively long oxidation reaction (24 h); similar observations were reported after
overnight incubation of Phanerochaete chrysosporium CDH with cellobiose [22].

Figure 3.NMR spectra of cellobiose (A) and xylobiose (B) oxidation. (A): From top to bottom are the spectra of cellobiose, cellobiose that was oxidized by
GOOX-VN, and cellobiose oxidized by Y300A; CB red. alpha and CB red. beta: H1 signals
due to reducing α-glucose and reducing β-glucose units of cellobiose, correspondingly;
CBA-H2 and CBA-H3: H2 and H3 signals of the cellobionate molecule. (B): From top to bottom are the spectra of untreated xylobiose and GOOX-VN oxidized xylobiose;
XB red. alpha and XB red. beta: H1 signals due to reducing α-xylose and reducing β-xylose
units of xylobiose, correspondingly; XBA: Overlapped signals of the xylobionate molecule
(H2 and H3 signals were not well separated from other signals). 10 mM cellobiose and
10 mM xylobiose were used in oxidation reactions.

ESI-MS/MS analyses also indicated enzymatic oxidation of cellotriose at the anomeric
carbon. In the positive ionization mode, the acidic fraction of oxidized cellotriose
only produced glycosidic bond cleavage fragments, generating B- and Y-ions (Figure 4A); cross ring cleavage fragmentation was not observed. Since neutral reducing oligosaccharides
usually form cross ring cleavage fragments from reducing ends if a sodium cation is
present [23,24], oxidation of the anomeric carbon seemed to change the fragmentation behaviour of
sodium cationized cellotriose. In the negative mode, B- and C-ions from glycosidic
bond cleavage were the most abundant fragment ions (Figure 4B). The molecular masses of Y- and Z-ions increased by 16 Da, compared to the unoxidized
control sample in our study (data not shown) or reported in the literature [25], supporting that the oxidation reaction occurred in the reducing glucose. Cross ring
cleavage fragmentation was also observed in the negative mode. For instance, a peak
at the m/z ratio of 383 was generated from oxidized cellotriose (m/z 519) by the loss of 136
Da from cross ring cleavage of the oxidized monosaccharide unit, leading to the formation
of a 2,4A3-ion (Figure 4B).

Additional, indirect evidence, from colorimetric assays, for the oxidation at C1 is
that no activity was detected on D-glucose derivatives lacking a C1 hydroxyl group,
including 1,5-anhydroglucitol (D-glucose with -H instead of -OH at C1) and methyl-β-D-glucopyranoside
(D-glucose with -OCH3 instead of -OH at C1).

Reconstructing the recombinant wild-type GOOX

The double substitution (V38A-N388S) was created in GOOX-VN to produce GOOX, the recombinant
wild type oxidase of S. strictum CBS 346.70, while the single N388S substitution was created to generate GOOX-V and
investigate the significance of V38.

The presence of the single A38V mutation increased the catalytic efficiency of GOOX-V
on all tested substrates compared to the wild-type enzyme; by contrast, the introduction
of both random mutations A38V and S388N (generating GOOX-VN) reduced enzyme activity
(Table 2). More specifically, comparisons of GOOX and GOOX-V revealed that the random mutation
of A38 to valine did not significantly change the Km but nearly doubled the kcat on all tested substrates. It was initially surprising that a mutation at position
38 affected enzyme activity since this position is close to the protein surface and
nearly 30 Å away from the oxidation site. Structural analysis of GOOX-T1 showed that
A38 is located on a flexible loop before the β2-sheet and it is close to D36 and E17,
which are predicted to coordinate one of the four zinc ions identified in GOOX-T1
crystals grown in the presence of 10 mM ZnSO4 (Figure 5) [20]. Since high quality crystals of GOOX-T1 were only formed in the presence of zinc
ions, it is possible that divalent ions coordinated by amino acids near A38 participate
in stabilizing the protein structure. Notably, the addition of 1 mM ZnCl2 slightly increases the specific activity of GOOX-T1, while 1 mM EDTA slightly reduces
the specific activity of the enzyme [10]. Since the β2-sheet, together with the β3-sheet and the β4-sheet, forms a P-loop
structure that participates in coordination [20], it is conceivable that the A38V substitution affects GOOX activity through an impact
on cofactor binding.

Figure 5.Structural analysis of GOOX-T1. (A): Metal binding sites of the FAD-binding domain (coloring by b-factor); A38 is close
to the zinc ion 2 bound to D36 and E17. (B): Residues for mutation in relation to the substrate analog, 5-amino-5-deoxy-cellobiono-1,5-lactam
(ABL); the movement of residues in the absence of ABL (PDB ID: 1ZR6, green) compared
with the presence of ABL (PDB ID: 2AXR, cyan) are indicated by arrows.

S388 is located in the β16-sheet, which forms part of the substrate-binding domain
(Figure 5). This position is close to a loop region formed by residues Y390 to N394, which
is absent in all of the 30 closest homologs of GOOX (analysis performed at http://consurf.tau.ac.il/webcite). The structure of GOOX-T1 bound by a substrate analogue, 5-amino-5-deoxycellobiono-1,5-lactam
(ABL) showed that the side chain of S388 rotates upon ABL binding to form a weak H-bond
with G349 [20], which is predicted to stabilize the β16-sheet. In this case, the comparatively large
side chain of asparagine could lead to steric destabilization of the protein. This
possibility is consistent with the comparatively low thermostability of GOOX-VN compared
to GOOX and GOOX-V (Figure 6), which could indirectly alleviate the beneficial affect of the A38V substitution.

Figure 6.Thermostability of GOOX, GOOX-V and GOOX-VN. Residual activity on 0.5 mM cellobiose, after a 60-min incubation at different temperatures
was measured at 37°C for 15 min at pH 8.0.

Impact of chain length and sugar composition on GOOX, GOOX-V and GOOX-VN activity

Mass spectrometric analysis of oxidized cello-oligosaccharides from cellobiose to
cellopentaose revealed a 16 Da increase in m/z values of the acidic fraction (Additional
file 3: Figure S3M-P) compared to the control, unoxidized samples (Additional file 3: Figure S3A-D), confirming that in all cases, the oxidation by GOOX-VN introduced
a single oxygen atom to all the oligomeric substrates. The oxidation of different
cello-oligosaccharides was efficient, but not complete at the tested concentrations,
as can be seen from small amount of unoxidized oligosaccharides detected in the neutral
fraction (Additional file 3: Figure S3I-L). Nevertheless, GOOX production of oxidized cello-oligosaccharides
might be an efficient way to generate oxidized carbohydrate standards to facilitate
the characterisation of the C1-oxidizing enzymes of families AA-9 and AA10.

H2O2-based colorimetric detection was then used to compare the catalytic efficiency of
GOOX, GOOX-VN and GOOX-V. Those analyses confirmed that the catalytic efficiencies
of these enzymes are over two orders of magnitude higher on oligomeric substrates
compared to corresponding monosaccharides (Table 2). Notably, however, among the oligomeric substrates, catalytic efficiencies with
cello-oligosaccharides and xylo-oligosaccharides decreased slightly with increasing
degree of polymerization, mainly due to increasing Km values (Table 2). This observation is consistent with earlier predictions of two binding subsites
in GOOX enzymes [20].

GOOX-VN, GOOX-V and GOOX effectively oxidized xylo-oligosaccharides as well as cello-oligosaccharides,
and GOOX displayed even higher catalytic efficiency on xylopentaose and xylohexaose
than on the corresponding cello-oligosaccharides (Table 2). The catalytic efficiency of GOOX and GOOX-V on a substituted xylo-oligosaccharide
(A3X) and a branched xylo-oligosaccharide (Ad2+3XX) was comparable to unsubstituted and unbranched substrates, indicating that these
sugar substitutions do not interfere with GOOX activity (Table 2). However, the catalytic efficiency of GOOX on U4m2XX was significantly reduced due to high Km values (Table 2), suggesting comparatively poor binding of anionic oligosaccharides by GOOX enzymes.
Regardless of a similarly low activity on xylose and two other monosaccharides: N-acetylglucosamine
and galactose [7], GOOX and GOOX-VN were not active on chitobiose, chitotriose, and galactobiose, suggesting
that substrate interaction at the -2 binding subsite plays an important role for substrate
specificity.

Key residues involved in substrate binding

Heuts et al. [26] reveal that by substituting one residue in the substrate recognition site of F. graminearum chito-oligosaccharide oxidase, the enzyme gains activity on gluco-oligosaccharides.
Accordingly, detailed site-directed mutagenesis was performed to study the contribution
of substrate-binding site residues on the substrate specificity and catalytic efficiency
of GOOX. Close examination of GOOX-T1 and GOOX-VN identified five amino acids in the
substrate-binding domain that were targeted for single mutation in GOOX-VN. Of the
chosen residues, Y72 and Q384 are at the -1 binding subsite, E247 and W351 are at
the -2 binding subsite and Q353 is between the two binding subsites (Figure 5). While most substitutions were to alanine, some were mutated to related residues
to evaluate the impact of amino acid size on enzyme activity. Since the catalytic
efficiency of GOOX-VN was not dramatically affected by oligosaccharide length (Table 2), oligosaccharides with up to 3 sugar units instead of 6 units were used to obtain
kinetic parameters for the mutant enzymes.

In all cases, Km values for the mutant enzymes were higher than GOOX-VN, consistent with the role
of each substituted residue in substrate binding (Table 3). For instance, the removal of either the hydroxyl group (Y72F) or the complete side
chain (Y72A) of an amino acid contributing to the -1 subsite increased the Km of corresponding enzymes, particularly on xylo-oligosaccharides (Table 3). Notably, Y72 interacts with the endocyclic O5 and can hydrogen-bond with OH6 of the cellobionolactone analog [20]. Huang et al. [14] observed that the H70A substitution in GOOX-T1, which removes a covalent linkage
to the FAD cofactor, also increases Km values. Given the close positioning of H70 and Y72, higher Km values in H70A mutants might result from indirect effects on Y72 positioning.

Table 3.Kinetics of GOOX-VN mutant enzymes on cello- and xylo-oligosaccharides

The impact of amino acid substitutions on catalytic rates was more varied. For instance,
kcat values of W351A on all cello-oligosaccharides were nearly three times higher than
corresponding values for GOOX-VN. Similarly, the kcat value of W351A on xylotriose was nearly triple that of GOOX-VN, even though activity
on xylose was not detectable (Table 3). Q384A lost activity on all substrates; however, the presence of asparagine at this
position could partially recover that loss, particularly on cello-oligosaccharides
(Table 3). Interestingly, the kcat of Q384N on glucose and cello-oligosaccharides was nearly doubled compared to GOOX-VN.
The distance between the O1 atom of the β anomer to the Oη of the catalytic base Y429 and to the Nϵ2 of Q384 is approximately 3.1 Å and 3.7 Å, respectively [20]. It is possible that the shorter asparagine side chain could improve the positioning
of C6 substrates relative to the catalytic base, thereby increasing kcat. Accordingly, the Q384N variant might be particularly useful when wanting to selectively
oxidize glucose and cello-oligosaccharides in mixtures containing xylo-oligosaccharides.

The mutation of Q353 to alanine eliminated enzymatic activity, which was not recovered
by replacing the alanine by asparagine (Table 3). Q353 forms two hydrogen bonds with the ABL substrate analog: one with the OH3 at the -1 subsite and the other with OH6 at the -2 subsite, which is the only direct protein-carbohydrate hydrogen bond in
the -2 subsite [20]. Structural analyses of GOOX-T1 bound by ABL also show that E247 shifts away from
the oxidation site, up to 4.6 Å from the unbound reference structure (Figure 5). The E247A variant displayed slightly reduced catalytic efficiency on all tested
substrates, generally resulting from increased Km values (Table 3). This result suggests that the predicted side chain movement at this position is
not crucial for enzyme activity.

The activity of GOOX and its two mutant enzymes GOOX-VN and GOOX-V was reduced at
comparatively high oligosaccharide concentrations, consistent with substrate inhibition.
A modified Hill model (Equation 2) [27] described the activity data better than the conventional uncompetitive substrate
inhibition model (Equation 1) (Figure 7A); therefore, inhibition kinetics parameters were calculated using Equation 2 (Table 4). Vi/Vmax ratios were also calculated since residual activities were measured for all substrates
and substrate concentrations tested, although to different levels. Most notably, Vi/Vmax values for cello-oligosaccharides were lower than for xylo-oligosaccharides, indicating
that the activity of all three GOOX variants was inhibited more severely by cello-oligosaccharides
than xylo-oligosaccharides (Figure 7B). Consistent with corresponding Km values (Table 2), Vi/Vmax values slightly increased with increasing cello-oligosaccharide chain length (Table 4). These inhibition data also indicate that GOOX can be a useful tool for glucose
as well as xylose oxidation at very high substrate concentrations.

Structural analyses suggest that non-productive substrate binding at high substrate
concentrations could be stabilized through stacking interactions with amino acid residues
above the Y300 and W351 -2 subsite (Additional file 4: Figure S4). This possibility of non-productive cooperative binding at the -2 subsite
is also consistent with alleviated inhibition observed upon Y300A [7] and W351A substitution (Table 4 and Figure 7A). Moreover, 1H NMR analyses were consistent with less inhibition of the Y300A enzyme by cellobiose
than GOOX-VN (Figure 3A). Specifically, the H1 signals due to reducing α-glucose and reducing β-glucose
units of cellobiose completely disappeared when 10 mM cellobiose was oxidized by Y300A
while their residual signals were detected when the same cellobiose concentration
was oxidized by GOOX-VN. These analyses suggest that W351A and Y300A mutant enzymes
might be ideal candidates for oxidizing otherwise inhibitory oligosaccharides when
present at high substrate concentrations.

Additional file 4: Figure S4. A potential non-productive binding subsite. (Left): A potential binding pocket (yellow
eclipse) above Y300 and W351, as seen in the GOOX-T1 structure with the presence of
ABL (PDB ID: 2AXR). (Right): A 45°-rotated view, the distance between two stacking
residues is 8.1 Å.

Conclusions

The double mutation of V38A-N388S in GOOX-VN to regenerate the recombinant wild type
oxidase of S. strictum CBS 346.70 confirmed that the reverse mutations do not explain the difference in
substrate preference between GOOX-VN and GOOX-T1. The current analysis also further
highlights that GOOX enzymes characterized to date are not specific to glucose-based
substrates, and instead show broad substrate specificity on a number of oligosaccharides
including cello- and xylo-oligosaccharides, as well as substituted and branched xylo-oligosaccharides.
The substrate promiscuity of GOOX, along with variants with higher catalytic activity
and lower substrate inhibition, broadens its applications in biomass processing at
high polysaccharide and oligosaccharide concentrations.

Methods

Materials

Sarocladium strictum type strain CBS 346.70 was obtained from the American Type Culture Collection (ATCC)
No.34717. Glucose, xylose, and cellobiose were purchased from Sigma (St. Louis, USA),
while other cello-oligosaccharides as well as xylo-oligosaccharides, chito-oligosaccharides
and galactobiose were purchased from Megazyme (Megazyme International, Ireland). Substituted
xylo-oligosaccharides including α-L-arabinofuranosyl-(1→3)-β-D-xylopyranosyl-(1→4)-D-xylose
(A3X), α-L-arabinofuranosyl-(1→2)-[α-L-arabinofuranosyl-(1→3)]-β-D-xylopyranosyl-(1→4)-β-D-xylopyranosyl-(1→4)-D-xylose
(Ad2+3XX) and 4-O-methyl-α-D-glucopyranosyl uronic acid-(1→2)-β-D-xylopyranosyl- (1→4)-β-D-xylopyranosyl-(1→4)-D-xylose
(U4m2XX) were prepared as previously described [28-30].

Site-directed mutagenesis

The QuikChange kit (Agilent Technologies, USA) and ten primer pairs (Additional file
5: Table S1) were used to separately introduce ten amino acid substitutions to GOOX-VN.
The GOOX-VN gene from S. strictum CBS 346.70 that was previously cloned into the pPICZαA expression vector [7] was used as the template for site-directed mutagenesis. Expression plasmids containing
the mutated gene were sequenced at the Centre for Applied Genomics (TCAG, the Hospital
for Sick Children).

Recombinant protein expression

Mutated plasmids were transformed into Pichia pastoris strains according to the manufacturer’s instructions (Life Technologies, USA). The
transformants were screened for protein expression by immuno-colony blot as previously
described [7] as well as using an overlay activity assay. Briefly, approximately 10 mL of the overlay
mixture (0.3% agarose, 2% cellobiose, 2 mM phenol, 0.4 mM 4-aminoantipyrine (4-AA)
and 15 U/mL horseradish peroxidase in 50 mM Tris–HCl pH 8.0) were applied over P. pastoris colonies that had been induced for 3 days by daily addition of 0.5% methanol. Following
30 min to 60 min of incubation at 37°C, transformants that expressed active forms
of the recombinant enzyme were identified by the formation of a pink halo around the
colony. Positive transformants were grown at 30°C and 250 rpm for 5 days, and 0.5%
methanol was added every 24 h to induce recombinant protein expression.

Culture supernatants were collected by centrifugation at 5,000 g for 10 min, and then
concentrated to approximately 15 mL using Jumbosep™ centrifugal concentrator units
(Pall Corporation, USA) before being passed through a HiTrap™ desalting column (GE
Healthcare, UK) using a BioLogic Duoflow FPLC system (Bio-Rad Laboratories, USA).
The protein fractions were loaded onto a GE HisTrap™ column (GE Healthcare, UK), washed
with the washing buffer (50 mM NaH2PO4, 300 mM NaCl, 20 mM imidazole, pH 8.0) and then eluted using the elution buffer,
which is the same as the washing buffer but with 250 mM imidazole. Eluted fractions
were replaced by 50 mM Tris–HCl (pH 8.0) using Vivaspin 20 concentration units (Sartorius,
Germany).

Confirmation of protein purity and identity

Protein concentrations were measured using the Bradford method (Bio-Rad Laboratories,
USA) and confirmed using SDS-PAGE densitometry, where the band density of GOOX and
the BSA reference protein were determined using ImageJ (http://rsbweb.nih.gov/ij/webcite). Retention of the FAD cofactor in mutant enzymes was assessed by verifying sample
absorbance at 350–700 nm using a Varian Cary 50 Bio UV–VIS spectrophotometer (Agilent
Technologies, USA). The presence of the FAD cofactor in intact protein samples was
further confirmed by running the enzyme samples using SDS-PAGE and incubating the
gel in 10% acetic acid for 10 min before visualization of fluorescent bands under
a hand-held Mineralight® UV lamp (UVP, USA). A second, identical SDS-PAGE gel was
treated with performic acid before the acetic acid treatment to check for increase
in fluorescence intensity [19] (note: extra caution is required when handling performic acid).

To confirm the introduction of single amino acid substitutions, protein samples were
exchanged to MilliQ water using 10 kDa Amicon filter units (EMD Millipore, USA), and
then 2000 pmol of each protein were processed using a Waters Pico-Tag System to evaluate
total amino acid composition (Advanced Protein Technology Centre, Hospital for Sick
Children, Toronto, Canada). Protein samples were also digested with modified sequencing-grade
trypsin (Promega, USA) and peptide sequences were obtained by tandem mass spectrometry
using an LTQ-XL™ mass spectrometer (Thermo Fisher Scientific, USA).

Enzymatic kinetics and thermostability

A 96-well chromogenic assay was used to measure hydrogen peroxide production [7,10]. Briefly, the production of H2O2 was coupled to the oxidation of 4-AA by horseradish peroxidase and measured continuously
at 500 nm and 37°C for 15 min. To determine specific activity, 16 nM of enzyme was
assayed with 0.5 mM oligosaccharides. Kinetic parameters were determined by using
16 nM of enzyme and a range of substrate concentrations: 0.05 - 300 mM glucose, 0.05
- 1200 mM xylose, 0.05 - 20 mM cellobiose, 0.01 - 10 mM cellotriose, xylobiose, and
xylotriose, 0.01 - 4 mM of longer cello- and xylo-oligosaccharides, 0.01 - 1 mM A3X and Ad2+3XX, and 0.04 - 0.4 mM U4m2XX. At least eight substrate concentrations in triplicates were assayed for each substrate,
and then kinetic parameters were calculated using the Michaelis–Menten equation of
GraphPad Prism5 software (GraphPad Software, USA). Substrate inhibition kinetics were
calculated using a conventional substrate inhibition equation (Equation 1) and a modified
Hill equation (Equation 2) [27]:

(1)

(2)

Where, Vi is the reaction velocity in the presence of inhibition and nH is the Hill coefficient.

To determine the temperature stabilities of enzyme variants, 16 nM of each enzyme
was incubated for 1 h at temperatures between 30 and 60°C, and residual activities
were measured using the conventional 4-AA chromogenic assay and 0.5 mM cellobiose.

Mass spectrometric analysis of oxidized products

Reaction mixtures containing 1 mM of cello-oligosaccharides, from cellobiose to cellohexaose,
and 160 nM GOOX-VN or GOOX-Y300A, in 50 mM Tris HCl (pH 8.0) were incubated overnight
at 37°C. To characterize oxidized products, 100 μL of each reaction mixture were diluted
in 900 μL of MilliQ-water, and diluted samples were purified and fractionated to neutral
and acidic oligosaccharides using a Hypersep porous graphitized carbon column (Thermo
Scientific, MA, USA), following the protocols of Packer et al. [31] and Chong et al. [32] with modifications. Neutral sugars were eluted using 40% acetonitrile, and mixture
of 50% acetonitrile and 0.05% TFA were used to elute acidic sugars. Collected fractions
were dried with nitrogen gas for 20 min and then freeze-dried overnight.

Mass spectrometric analyses were performed using an Agilent XCT Plus model ion trap
mass spectrometer (Agilent Technologies, Waldbronn, Germany) equipped with an electrospray
source. For ESI-MS and ESI-MS/MS analyses, freeze dried samples were dissolved in
20 μL of MilliQ-water, and 6 μL of each sample was diluted in 100 μL of methanol–water-formic
acid solvent (50:49:1 (v:v:v)). Sample solutions were introduced into the ES source
at a flow rate of 5 μL/min via a syringe pump. The drying gas temperature was set
to 325°C; drying gas flow was 4 L/min; the nebulizer pressure was 15 psi, and the
ES capillary voltage was set to 3164 V. Ions were collected in the m/z range of 50
to 1000. ESI-MS/MS analyses were performed in both positive and negative ionization
modes. Fragmentation amplitude was set to 0.60 V in the positive mode and 0.80 V in
the negative mode, and the precursor ion isolation width was set to 1.0 m/z and 1.5
m/z, respectively.

Competing interests

The authors declare there are no competing interests.

Authors’ contributions

TVV, JS, MT and ERM designed the experiments. TVV, AHV, MF and MJ performed the experiments
and analyzed the data. All authors discussed the results and implications and commented
on the manuscript at all stages. All authors read and approved the final manuscript.

Acknowledgements

Funding for this research was provided by the Government of Ontario for the project
“Forest FAB: Applied Genomics for Functionalized Fibre and Biochemicals” (ORF-RE-05-005),
grants from the Natural Sciences and Engineering Research Council, and a FiDiPro Fellowship
to ERM from the Finnish Funding Agency for Technology and Innovation (Tekes). This
study was also funded by the Academy of Finland for the ENOX project (Number 252183)
to MT and (Number 252827) to ERM.